(1:1000 = 1 mg/mL)
Conversion of coarse fibrillation is based on the use of cardiac massage as described above, or if the clinic has access to defibrillation devices then the use of these externally at 2–10 J/kg (starting at low energies and increasing if no response is achieved) may be performed (Costello, 2004). Greater success is achieved with defibrillation devices if three initial countershocks are applied at low energies.
If severe bradycardia is detected, glycopyrrolate should be used (see Table 8.1) in preference to atropine as 60% of domestic rabbits possess serum atropinesterases making atropine less effective (Okerman, 1994).
Lidocaine may be administered intratracheally or intravenously if ventricular arrhythmias such as ventricular premature complexes leading to ventricular tachycardia occur (see Table 8.1 for dose). However, lidocaine should not be used in cases of AV block or severe bradycardia (DeFrancesco, 2000), which are more commonly seen in rabbits.
Cardiomyopathy and valvular insufficiency with resultant congestive heart failure are also seen in rabbits as is atherosclerosis. Treatment of congestive heart failure initially depends on the use of diuretics such as furosemide at 1–4 mg/kg intravenously, repeated every 4–6 hours as required. ACE inhibitors have been used in rabbits but they are more susceptible to hypotensive side effects than cats or dogs. Therefore, reduced dosages and regular monitoring of the systolic blood pressure using non-invasive techniques devised for cats are advisable (Girling, 2003).
Diagnostic procedures
ECG (E)
As mentioned above, should cardiac arrest or arrhythmias be detected, ECG leads may be applied as in cats and dogs and the trace assessed. See Table 8.2 for some normal values for rabbits.
Table 8.2 Normal ECG values (lead II) for healthy rabbits.
| Parameter | Normal result | Notes |
| P wave height | 0.1–0.15 mV | Deflection is low or negative in lead I and always positive in leads II and III |
| P wave duration | 0.03–0.04 s | |
| P-R interval | 0.05–0.1 s | |
| QRS complex duration | 0.015–0.04 s | |
| R wave amplitude | 0.3–0.39 mV | |
| Q-T interval | 0.08–0.16 s | Change of deflection of the T-wave from positive to negative or vice versa indicates myocardial hypoxia as with cats and dogs |
| Source: From Kozma et al. (1974) and Huston and Quesenberry (2004). | ||
Radiography
This is essential where acute gastrointestinal signs are present, such as bloat or visceral pain, or where evidence of limb fractures, paresis or paralysis is present and should be performed when clinically safe to do so as part of a full clinical work-up. In this chapter there is not enough time to go into detail regarding diagnostic imaging but the following conditions may be associated with radiographically visible changes.
Abdominal pain may be associated with a number of conditions including renal calculi, obstructive and non-obstructive intestinal ileus and peritonitis. Spinal luxations, fractures and dislocations may also be identified radiographically. Gradation of severity may be made as with cats and dogs. Acute treatment with short-acting intravenous corticosteroids (e.g. dexamethasone, see Table 8.1) and immobilisation with analgesia are advised for spinal trauma with concurrent paresis or paralysis as for other small mammals. Surgery may then be performed for collapsed discs or fractures as for cats and dogs as required.
Ultrasound
This may be of use where renal, bladder, cardiac and hepatic diseases are concerned. Cardiomyopathies (dilated) and atherosclerosis have been reported. Hepatic lipidosis is common in rabbits. Urinary bladders often contain large amounts of silt which create a snow-storm effect.
Blood testing
Routine haematology and biochemistry assessment should be made – values may be found in standard rabbit textbooks. On a critical care level, blood lactate levels may also be measured in rabbits with levels above 10 mmol/L being viewed as abnormal (Lichtenberger & Ko, 2007).
Fluid therapy (F)
Hypovolemic shock
If the rabbit is dehydrated or hypovolemic, shock doses of fluids should be administered. It should be noted that fluids should be avoided post resuscitation in cases of cardiovascular arrest where there is no hypovolemia/
dehydration prior to the arrest as these fluids may decrease myocardial perfusion pressures and diminish overall nutrient delivery through the cerebral and coronary vasculature (Cole et al., 2002). Rabbits tend to more closely mimic cats in hypovolemic shock with the decompensatory stage being seen without compensatory shock, i.e. evidence of bradycardia, hypotension and hypothermia. In these cases a slow intravenous bolus of fluids such as hypertonic saline 7.2–7.5% at 3 mL/kg to draw fluid rapidly into the circulation is advised (Lichtenberger & Lennox, 2010). This can be maintained by follow-up administration of Hetastarch (3 mL/kg) over 10 minutes. The patient should be warmed, and crystalloids at 3–4 mL/kg/h should be administered. It is important to measure systolic blood pressure during this procedure. Once the rabbit has been warmed, the aim is to get the systolic blood pressure above 90 mm Hg. This may require further boluses of isotonic crystalloids (10 mL/kg) with Hetastarch (5 mL/kg). Once normovolaemia has been achieved, replacement of fluid deficits may be started (remember maintenance values for rabbits are 80–100 mL/kg per day).
General fluid administration
Oral
Not such a good route for seriously debilitated animals but useful for those with naso-oesophageal feeding tubes in place. Again it may be useful for mild cases of dehydration where owners wish to home treat their pet. This route though is restricted to small volumes with a maximum of 10 mL/kg at any one time administered. In practice it may be possible to administer much less than this.
Subcutaneous
The scruff area or lateral thorax makes ideal sites. This is a good technique for routine post-operative administration of fluids for longer recovery patients undergoing minor surgical procedures such as spaying or castration. It is possible to give a maximum of 30–60 mL split into two or more sites depending on the size of rabbit at one time.
Intraperitoneal
To perform this it is necessary to tilt the rabbit’s head downwards whilst it is in dorsal recumbency to allow the gut contents to fall out of the injection zone. The needle is inserted in the caudal right quadrant of the ventral abdomen. It is inserted just through the abdominal wall and the plunger drawn back on the syringe to ensure that no puncture of the bladder or gut has occurred. A maximum volume of 20–30 mL may be given at one time depending on rabbit size. Previous notes regarding concurrent respiratory or cardiovascular disease should be considered. If positioned correctly, there should be no resistance to injection.
Intravenous
Venous access is relatively straightforward in the rabbit. The marginal ear vein, jugular vein, cephalic vein and lateral saphenous vein can all be used for intravenous catheter placement. Long-term (>2–3 days) use of the marginal ear vein may, however, cause sloughing of the ear tip, although in the author’s experience this is relatively rare with careful venipuncture technique. Use of a topical local anaesthetic cream is recommended prior to placement. In addition, if a sedative such as Hypnorm® (VetaPharma UK Ltd.) is used (a fentanyl/fluanisone combination neuroleptanalgesic drug), peripheral vasodilation is common facilitating ear vein catheter placement. The fentanyl portion of this drug may be reversed using the partial opioid agonists – buprenorphine (0.01–0.05 mg/kg) or butorphanol (0.1–0.5 mg/kg). The jugular vein may be difficult to access, particularly in does where there is a pronounced ruff of skin and fat deposits. In addition it forms the main venous drainage for the eye, and thrombus formation may lead to periocular swelling. An Elizabethan collar can be used to prevent the rabbit from chewing or removing the catheter, although this will prevent caecotrophy and care should be taken to ensure the rabbit is still managing to eat, otherwise assisted feeding (see below) should be instituted.
Intraosseous
Intraosseous catheters may be placed into the proximal femur, in the trochanteric fossa, in a parallel direction to the long axis of the femur. Use an 18–23 gauge, 1–1.5 inch spinal or hypodermic needle. Analgesia should be employed (see Chapter 3) whenever placing an intraosseous catheter as should prophylactic antibiosis such as enrofloxacin (Baytril 2.5% Bayer – licensed for use in rabbits).
Intraosseous and intravenous fluid administration should be accurately titrated using syringe drivers rather than relying on drip sets since even a small error in fluid administration may be proportionally more significant considering the small size of many rabbits.
Other medications and supportive nutrition
Many rabbits presented as an acute emergency either have already or frequently go on to develop gastrointestinal stasis. Providing obstructive causes have been ruled out; the use of prokinetic medications, such as cisapride, metoclopramide and ranitidine, should be performed (see Table 8.3). Ranitidine acts to reduce acidity in the stomach, which is beneficial as many rabbits with gastrointestinal stasis have punctate ulceration of the stomach lining. In humans it also has prokinetic effects encouraging emptying of the stomach and some increased motility of the small intestine, which seems to be synergised by metoclopramide; this appears clinically to be the case in rabbits in this author’s and others’ opinion.
Table 8.3 Gut motility enhancing drugs for rabbits.
| Drug | Dose rate | Frequency of dosing and notes |
| Cisapride | 0.5 mg/kg PO | 12 hourly |
| Metoclopramide | 0.5 mg/kg SC | 8–12 hourly |
| Ranitidine | 2–5 mg/kg PO | 12 hourly (in combination with metoclopramide acts to promote motility as well as reducing acidity) |
| Note that none are licensed for use in rabbits. | ||
Assisted feeding should be carried out in conjunction with the use of prokinetics. This can start off with easily absorbed essential sugars and amino acids (e.g. Critical Care Formula®, Vetark Professional) either syringed into the mouth or delivered via a naso-oesophageal tube. As the rabbit improves clinically, this should be stepped up to use proprietary critical feeding formulas such as Science Recovery® and RecoveryPlus® (Supreme Petfoods) or Critical Care for Herbivores® (Oxbow Pet Products) or vegetable-based baby foods (lactose-free varieties). The disadvantage of baby foods is that they do not contain fibre and thus have little or no prokinetic activity, although they do provide nutrients in an easily digestible form.
The levels of energy required for a debilitated rabbit should approach that calculated for growing-to-lactating rabbits using the formula – MER = k × (wt [kg])0.75, where k = 200 for growth and 300 for lactation (Carpenter & Kolmstetter, 2000). Therefore for debilitation, the following daily energy requirement may be used:
MER = 250 × (wt [kg])0.75
To re-populate the intestinal flora, transfaunation of caecotrophs from a healthy rabbit may aid the return of normal bowel function. The use of commercial probiotics designed for rabbits has also been advocated, and reduces the risk of transferring potential parasites and other agents to the debilitated patient.
Naso-oesophageal tube in rabbits
A naso-oesophageal tube (as the tubing must not allow reflux of acid stomach contents into the oesophagus) is placed after first spraying the nose with lidocaine spray and inserting the 3–4 French tube which has been pre-measured from the extended nose to the seventh rib. Sterile water should be flushed through the tube before and after feeding to ensure it is correctly placed and does not become blocked. The tube may then be glued, taped or sutured to the dorsal aspect of the head and a bung inserted when not in use. It may be necessary to put an Elizabethan collar on the rabbit to prevent removal.
EMERGENCY CARE OF OTHER SMALL MAMMALS
Emergency airway access and ventilation (A and B)
Providing a direct airway may be very difficult in rodents. In ferrets and omnivorous marsupials, intubation is similar to small cats. Access to the epiglottis is difficult owing to the nature of rodent anatomy, whereby the soft palate is locked around the epiglottis making its access via the mouth very difficult.
If the rodent is still breathing, placement in a small container and piping in oxygen may be all that is necessary.
If the small mammal is not breathing, then any of the following three techniques may be attempted.
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